On-Chip Fabrication of a Cell-Derived Extracellular Matrix Sheet

Nov 13, 2017 - fibronectin fibrous architecture were maintained on the surface of the porous membrane of the microfluidic device after ... fluidic sys...
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On-Chip Fabrication of a Cell-Derived Extracellular Matrix Sheet Yoonmi Hong,† Ilkyoo Koh,† Kwideok Park,‡ and Pilnam Kim*,† †

Department of Bio and Brain Engineering, Korea Advanced Institute of Science and Technology, Daejeon 34141, Republic of Korea Center for Biomaterials, Korea Institute of Science and Technology, Seoul 02792, Republic of Korea

‡

ABSTRACT: The extracellular matrix (ECM) provides physical and chemical support to the surrounding cells. During cell growth, ECM secretion and network formation influence cell morphology, cell adhesion, cell-to-cell interactions, and cell migration. Microfluidics-based cell culture systems are limited by the integration of structural ECM into the device. We report the development of a cell-derived ECM-incorporated microfluidic device that can provide structural characteristics and biochemical components of cell-derived ECM. Using an on-chip decellularization process, we constructed an ECM sheet, secreted and deposited from monolayer-cultured mouse embryonic fibroblasts (NIH/3T3), inside the microfluidic device. ECM components (including collagens, fibronectin, laminin, and elastin) and mesh-type fibronectin fibrous architecture were maintained on the surface of the porous membrane of the microfluidic device after decellularization. To verify the usability of the fibroblast-derived ECM sheet integrated microfluidic device in a cell culture platform, we tested the recellularization of human umbilical vein endothelial cells (HUVEC) and analyzed HUVEC−ECM and HUVEC−HUVEC interactions. On the ECM sheet, HUVECs exhibited morphologies and focal adhesion features that were markedly different from those of other groups. We then explored the effect of the ECM sheet on HUVEC mechanosensitivity. An increase in fluid shear stresses led to focal adhesion and the polymerization and reorganization of HUVEC adherens junctions, similar to natural junctional development, whereas the control group exhibited stimuli-insensitive behaviors. We conclude that the decellularized ECM sheet-incorporated microfluidic device provides an in vivo-like physical and biochemical ECM microenvironment for microfluidics-based cell culture. KEYWORDS: decellularization, microfluidics-based cell culture, fibroblast-derived extracellular matrix (ECM) sheet

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INTRODUCTION Microfluidic devices have been used as a platform for multiple applications, including in biosensors,1 droplet-based microfluidics,2,3 enzymatic analysis,4 protein chips,5 single-cell biology, and organ-on-chips.6−9 The culture of cells in microfluidic devices is becoming a particularly popular tool within experimental bioengineering. One of the advantages of the microfluidics-based cell culture system is the ability to mimic the natural microenvironment of cells, as a precursor to modulating their function.10 To implement an in vivo microenvironment, researchers have incorporated various stimuli into the microfluidic systems, including cyclic stretching,11 fluid shear stress,12 or an electrical field.13 For this purpose, it is necessary to attach the cells stably, providing a physiologically relevant surface within the microfluidic system. It is common to treat the cell-adhesive substrate with an extracellular matrix (ECM) associated with the microenvironment of the specific cells,14 with a chemical substance to be implicated in a cellular attachment via integrins,15 or with a molecule binding to the charged cell bilayer membrane,16,17 thereby enhancing cell growth. These procedures in the substrate do not sufficiently provide cell−substrate interactions and are not suitable for the simultaneous effects providing © XXXX American Chemical Society

complex ECM components and architecture, such as basement membrane (BM) or interstitial tissue. Recently, cell-derived ECM, which is obtained via a decellularization procedure, has been suggested as a physiologically relevant ECM microenvironment for tissue engineering.18−21 Cell-derived ECM is composed of cell-secreted ECM and can leave the physical network of cross-linked ECM.22,23 Indeed, the native substances of ECM have advantages in terms of matrix fibrils, porosity, permeability, and mechanical stability, which play important roles in the promotion of cell function and in the regulation of tissue regeneration.24 Cell-derived ECM also avoids host immune response to reseeded cells, relative to organ-derived ECM.25 Several studies have demonstrated its various applications, based on its capacity to regulate cell attachment and migration on cell-derived ECM.25,26 In this study, we developed an on-chip decellularization and recellularization process, and incorporate a cell-derived ECM sheet and the microfluidic device. The microfluidic device was composed of two layers of compartments (microchannel and static well) and a microporous membrane. To obtain the cell-derived ECM sheet, mouse embryonic fibroblasts (NIH/ Received: August 24, 2017 Accepted: November 13, 2017 Published: November 13, 2017 A

DOI: 10.1021/acsbiomaterials.7b00613 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering

Figure 1. (A) Illustration of the chip fabrication process. The microfluidic device was composed of a single microchannel, polyester microporous membrane, and parabolic well. (B) Decellularization and recellularization overview. Mouse embryonic fibroblasts (NIH/3T3) were cultured in the microchannel for 1 week. A fibroblast-derived extracellular matrix (FB-dECM) was obtained by decellularization of monolayered fibroblasts treated with a surfactant cocktail. Human umbilical vein endothelial cells (HUVEC) were recellularized on FB-dECM until full confluency for endothelialization was attained. To create the microfluidic chip, we irreversibly assembled the single microchannel, microporous membrane, and PDMS well after oxygen plasma treatment (Cute, Femto Science Inc.) for 1 min. The microfluidic chip sandwich assembled with the single microchannel, microporous membrane, and PDMS well (Figure 1A). Decellularization and Recellularization. The NIH/3T3 cell line was seeded at 1.5 × 105 cells/mL and cultured until full confluence on the microfluidic chip was attained; the medium was changed every day. To culture NIH/3T3 cells and HUVECs, we supplie the mediad with a gravity flow using 200 μL yellow pipet tips on the inlet and outlet reservoirs. To perform the decellularization and microfluidic experiments, we connected the microfluidic chip to a syringe pump. A surfactant cocktail was prepared for decellularization using a mixture of 0.1% sodium dodecyl sulfate (SDS) (Sigma) and 0.1% triton X-100 (Sigma) in deionized water, which was stirred at 50 rpm overnight. The surfactant cocktail was warmed in a water bath at 37 °C before use. After 6−7 days of culture (until full confluence), fibroblasts were washed with Dulbecco’s phosphate-buffered saline (DPBS, Welgene Inc.) and the decellularization cocktail was exposed to a 1-dyn/cm2 shear stress for a few seconds using a syringe pump. After decellularization, the remaining fibroblast-derived ECM was washed with 0.001% Poly-L-Lysine (PLL, MW = 70 000−150 000, Sigma) in DPBS. HUVECs were reseeded with 1.5 × 105 cells/mL on the decellularized microfluidic chip, and cultured until full confluence was attained. Various ECM Coating Used on the Microfluidic Channel Substrate. To determine cell-matrix effects on cell attachment and focal adhesion formation, we compared three types of ECM. We used 100 μg/mL of fibronectin from bovine plasma (Sigma), 100 μg/mL of mouse Matrigel (Corning Inc.), and 100 μg/mL of human placental laminin (Sigma) to coat the microfluidic channel at 37 °C and 5% CO2 in an incubator overnight. Following aspiration of the ECM solution, HUVECs were seeded on the microfluidic chip. At full confluence, the HUVECs were prepared for immunostaining and real-time qualitative polymerase chain reaction (RT-qPCR). Immunofluorescence Staining. Cells were gently washed with DPBS and fixed with 4% paraformaldehyde (Biosesang) for 15 min at room temperature. After fixation, the microporous membrane removed from the microfluidic chip by cutting the PDMS with scissors. The cells were then blocked with 1% bovine serum albumin (BSA) (Sigma) in DPBS for 5 min and permeabilized with 0.3% triton X-100 (Sigma) in DPBS for 10 min. Primary antibodies were incubated for 1 h, and fluorescent-conjugated primary and secondary

3T3) were cultured on the microporous membrane within the microchannel for 1 week, and the monolayered fibroblasts were then treated with a decellularization cocktail. We verified the architecture of the fibroblast-derived ECM sheet by immunofluorescence staining of the major ECM proteins. Human umbilical vein endothelial cells (HUVECs) were then sequentially reseeded on top of the ECM sheet. We characterized the HUVEC−ECM interaction and the adherens junctions of the reseeded HUVEC. Furthermore, shear stressinduced junctional polymerization and adhesion reorganization were observed through immunostaining. These results demonstrate that the combination of a cell-derived ECM sheet-based culture system with application of shear stress is appropriate for endothelialization within a microfluidic device for tissue engineering and other biomedical applications.

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MATERIALS AND METHODS

Cell Culture. We performed decellularization using a mouse embryonic fibroblast cell line (NIH/3T3) purchased from the Korean Cell Line Bank (KCLB). NIH/3T3 cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) (Welgene Inc.) supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin (Welgene Inc.). HUVEC cells were cultured in EGM BulletKIT medium (LONZA). Both cell lines were cultured at 37 °C and 5% CO2 in an incubator. Microfluidic Chip Fabrication. A single microchannel (width: 1 mm, length: 15 mm, height: 100 μm) was prepared on a silicon master that had been prepared by deep reactive ion etching (DRIE). To fabricate the top layer of the polydimethylsiloxane (PDMS) (Sylgard 184 Silicon Elastomer Kit, Dow Corning) channel, a PDMS elastomer base and curing agent (10:1 w/w) mixture was poured onto the silicon master and baked in a convection oven at 70 °C for 2 h. Bare PDMS was prepared using the same procedure for the bottom layer. A single microchannel was prepared by peeling out from the silicon master; the inlet and outlet were created by punching a 1 mm hole into the PDMS replica. A parabolic well (width: 2 mm, length: 1 cm, height: 2 mm) was punched into a bare PDMS slip to prepare the bottom reservoir. A random microporous membrane (width: 4 mm, length: 20 mm) was cut with scissors from a Transwell insert membrane (0.4 μm of the pore, 10-μm thickness polyethylene terephthalate (PET)) (Falcon). B

DOI: 10.1021/acsbiomaterials.7b00613 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering

Figure 2. (A) Characterization of fibroblast-derived ECM. Representative images include immunofluorescent staining for ECM proteins (fibronectin, collagen I, collagen IV, elastin, and laminin) from mouse embryonic fibroblast (NIH/3T3) secretion and the decellularized ECM sheet. Blue indicates nuclei (scale bar = 50 μm).

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antibodies were incubated for 40 min at room temperature, and they washed three times with DPBS for (5 min each). All antibodies are described below. We incubated 4′,6-diamidino-2-phenylindole (DAPI, Sigma) for 5 min at room temperature; cells were then covered with a mounting reagent containing Prolong Gold Antifade Reagent (Invitrogen). The fluorescent images were captured under an inverted fluorescent microscope (Leica 4000B, Germany), and a confocal laser microscope (Nikon Ti-E, Japan). Conjugated primary antibodies included tetramethylrhodamine B isothiocyanate (TRITC) phalloidin (1:200 in DPBS) (Sigma), FITC antivinculin in mouse (1:100 in DPBS, Sigma), FITC anti- β-catenin (1:100 in DPBS), and DAPI (1:5000 in DPBS, Sigma). Primary antibodies included rabbit anti-fibronectin (1:100 in DPBS, Sigma), rabbit anti-collagen I (1:100 in DPBS, Sigma), rabbit anticollagen IV (1:100 in DPBS, Abcam), rabbit anti-laminin (1:100 in DPBS, Sigma), rabbit anti-elastin (1:100 in DPBS, Abcam), and rabbit anti-VE cadherin (1:100 in DPBS, Cell Signaling). Secondary antibodies included goat anti-rabbit IgG-FITC (1:80 in DPBS, Sigma), and goat anti-rabbit IgG-TRITC (1:100 in DPBS, Sigma). RNA Preparation and RT-qPCR. Total RNA was isolated from cells using TRIZOL reagent (Thermo Scientific). RNA samples (1 μg) were reverse transcribed using an RT-PCR kit (Toyobo), according to the manufacturer’s instructions. The following primers were used for gene amplification; VE-cadherin, forward 5′-CAGCCCAAAGTGTGTGAGAA-3′ reverse 5′-TGTGATGTTGGCCGTGTTAT-3′, β-catenin, forward 5′-AAAATGGCAGTGCGTTTAG-3′ reverse 5′TTTGAAGGCAGTCTGTCGTA-3′, GAPDH, forward 5′-GTATGACAACAGCCTTCAAGAT-3′ reverse 5′-AGTCCTTCCACGATACCAAA-3′. All primer sets were optimized to ensure efficient amplification of a single product before use. We performed qRTPCR using the SYBR kit according to the manufacturer’s instructions (Toyobo) in the CFX Manager (Biorad, Hercules, CA, USA). Relative mRNA levels were calculated using the 2-ΔΔCt comparative method. Statistical significance of gene levels in cells was determined using Student’s t test, at P < 0.05. Flow Experiments. After HUVECs were reseeded on the ECM sheet and cultured for 10 days until full confluency was attained. HUVECs were exposed to laminar flows of 0.5, 1, and 5 dyn/cm2 for 2 h in EGM BulletKIT medium. The flow rate of the rectangular microchannel was calculated using the following equation:

τ=

RESULTS AND DISCUSSION On-Chip Decellularization Procedure. Figure 1A shows a schematic diagram of the fabrication of PDMS microfluidic device composed of a cell-derived ECM sheet. The microfluidic system consists of a single microfluidic channel, microporous membrane, and substrate with a parabolic well. Figure 1B shows the procedure of decellularization of monolayered NIH/ 3T3 fibroblasts (FB) and recellularization of HUVECs on top of the ECM sheet. Fibroblasts were cultured on the microporous membrane within the microchannel, and the ECM sheet was obtained by decellularization of the monolayered fibroblasts. After HUVEC endothelialization, the single microfluidic channel provides a lumen-like physiology, where endothelial cells are exposed to a fluid flow (Figure 1B). The ECM sheet on the microporous membrane acts as a BMlike microenvironment, i.e., an ECM layer separating endothelial and connective tissue layers in blood vessels. Characterization of Fibroblast-Derived ECM in Chip. During culturing, the fibroblasts deposit various ECMs including abundant fibronectin, several types of collagen, and laminin, which we observed via immunofluorescence staining (Figure 2). At full fibroblasts confluence, fibronectin fibrils were assembled and interlaced with fiber bundles. The fibronectin fiber structure displayed an irregular arrangement without network formation. Collagen I, collagen IV, and laminin were observed as parabolic features surroundings nuclei in cell bodies, whereas elastin was dispersed in the form of particles. After decellularization, most ECM components remained, but their architectures were destroyed due to the loss of cellECM integrity. Cells lose volume after treatment with surfactant mixture, i.e., the decellularization cocktail, leaving only the ECM sheet on the surface of the microfluidic chip. The complex architectures of fibronectin fibers were preserved with high porosity. Collagen I and IV were un-cross-linked, whereas there was no significant change in elastin. Laminin was dispersed, without macroscopic architecture. Presumably, laminin and other ECM fragments were swept out in the decellularization cocktail flow. Characterization of Adhesion between ECM Sheet and Reseeded HUVECs. To investigate the anchoring properties of HUVECs on the decellularized ECM sheet, we observed actin filaments and localized vinculin via immunostaining at 6, 24, and 48 h after seeding HUVEC seeding. We compared the HUVEC-decellularized ECM sheet interaction with those from previous studies using conventional

6μQ bh2

where τ is the flow rate (dyn/cm2), μ is the medium viscosity (gm/ cm/s), Q is the volumetric flow rate (cm3/s), b is the channel width, and h is the channel height. C

DOI: 10.1021/acsbiomaterials.7b00613 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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Figure 3. Morphological characterization of recellularized HUVECs on an ECM sheet in a microfluidic channel coated with 50 μg/mL fibronectin, 50 μg/mL laminin, 50 μg/mL Matrigel, and in a control washed in Dulbecco’s phosphate-buffered saline (DPBS) at 6, 24, and 48 h after seeding. (A) HUVEC F-actin (red) and nuclei (blue). Scale bar = 100 μm. (B) Quantification of the area of a single cell on an ECM sheet and a different ECM, 6 h later after seeding. n ≥ 40 cells, ** P < 0.005. (C) Local images of HUVEC attachment and spreading on the ECM sheet, F-actin (red), vinculin (green), and nuclei (blue) of HUVEC were immunostained. White box = 10 μm.

approaches, including ECM-coated surfaces (e.g., fibronectin and laminin), Matrigel, and a nontreated surface as a control. During the first 6 h of the initial cell culture period, the HUVECs were aligned along the flow direction during seeding (Figure 3A). The cells spontaneously attached to various ECM protein-coated surfaces and control. The cell bodies were spindle-shaped, and the HUVEC adhesion sites were localized in a dot-shape on the ECM sheet (Figure 3C). In contrast, the localized adhesion sites of different ECM-coated surfaces and the control were not detected by vinculin staining. The vastly expanded cells were observed on fibronectin- and Matrigelcoated surfaces. There were fewer attached cells on the laminincoated surface than on the other surfaces. We observed cell spreading in a triangular shape on the nontreated surface. An analysis of the HUVEC spreading area 6 h after seeding showed that the smallest value of (700 μm2) was associated with the ECM sheet. Spreading on the fibronectin- and Matrigel-coated surfaces were observed to be approximately triple and double the area of cell spreading on the ECM sheet, respectively (Figure 2B). The endothelial cell area typically occupies approximately 300−1200 μm2, which resembles the size of in vivo endothelial cells.26

After the initial growth period, within 24 h, the results of phalloidin staining indicates polymerization of F-actins was relatively high rates on both ECM sheet and laminin-coated surface (Figure 3A). On the laminin-coated surface, the cells exhibited highly clustered thick and abundant with more organized actin fibers on the cell membrane. In parallel, as a result of vinculin staining, the dense and localized focal adhesion was colocalized at the cell bodies on both ECM sheet and laminin-coated surfaces (Figure 3C). In the HUVEC on the fibronectin- and Matrigel-coated surface, the polymerization of F-actins was observed shorter fibers than that of the control group. After 24 h in culture, on the ECM sheet, it is difficult to distinguish the boundaries between the cells, but it showed a polymerized F-actin development that could roughly predict the morphology of the one cell (Figure 3A). The vinculin localization was observed in two types as dot- and line-shaped pattern on the ECM sheet. On the different ECM-coated surface and the control, high polymerization of F-actin was observed at the cell perimeters and randomly distributed Factin was developed at the interior of cells. D

DOI: 10.1021/acsbiomaterials.7b00613 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering

Figure 4. (A) Immunofluorescent image of recellularized HUVEC adherens junction expression on ECM sheet. Scale bar = 50 μm. Red indicates VE-cadherin, blue indicates β-catenin and nuclei. (B) Gene expression analysis of HUVECs on dECM (VE-cadherin, β − catenin). n ≥ 4, mean ± s.d. * P < 0.05.

Figure 5. Organization of HUVEC cells focal adhesion and cell junction under three different fluidic conditions. Green, red, and blue indicate vinculin, VE-cadherin, and nuclei, respectively. (a−c) HUVEC reseeded on the ECM sheet; (d−f) HUVECs on the control (microporous membrane). Scale bar = 20 μm; white box = 20 μm × 20 μm in magnified view. (B) The length of focalized vinculin of HUVEC on cell-derived ECM sheet. n ≥ 50, mean ± s.d. ***, ### P < 0.001.

Based on our results, we confirmed that the structurally defined ECM sheet could regulate the cytoskeleton arrangement and focal adhesion of endothelial cells in the initial stage of anchoring. The effect of the ECM sheet on the HUVEC cytoskeletal F-actin could be categorized as both compositional and structural. Initially, the HUVEC may recognize the ECM sheet as a three-dimensional (3D) matrix, consistent with previous observations.27 Moreover, the fibrils and the bundles of the ECM sheet could provide a nano- or microsized structural stimulus for HUVEC attachment, facilitating spindle and elongated morphologies, with localized focal adhesion sites.27,28

In addition to the structural effects, the biochemical effects of the ECM sheet on the HUVECs was observed in the biochemical conditions of fibronectin, laminin, and Matrigel (collagen IV and laminin-rich matrix); in this case, the ECM sheet contained BM constituents and fibronectin, which provides a physiologically relevant cell-ECM interaction, unlike the ECM protein-coated surface. Localization of the Adherens Junction on the ECM Sheet. We next explored the effects of the ECM sheet on the stability and maturation of recellularized endothelial cells. The integrity of the cell−cell junctions of endothelial cells is critical for maintaining a stabilized state.29 To quantify the protein E

DOI: 10.1021/acsbiomaterials.7b00613 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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CONCLUSION We demonstrated an effective culture system to create the physiological conditions of VE cells by obtaining fibroblastderived ECM in a microfluidic chip. After decellularization in a microfluidic channel, we observed the compositional heterogeneity and nano- to the microstructural architecture of FBdECM were observed, verified that the effects of HUVECs on the development of the cytoskeleton, focal adhesion, and adherens junctions on an ECM sheet. Furthermore, we confirmed that HUVECs recellularized on an ECM sheet exhibited long lines of focal adhesion under fluid shear stress, which indicates that endothelial cells can ensure fast blood flow through strongly developed cell−cell and cell−ECM interactions. Thus, the on-chip decellularization method that we present may be useful in a variety of tissue engineering, organon-a-chip, and within-microchannel endothelialization applications.

expression of the adherens junction, we analyzed the immunofluorescence-stained images using the image processing and analysis software, NIH ImageJ (Figure 4). The regulation of the adherens junction was visualized using vascular endothelial VE-cadherin and β-catenin. The localization of VE-cadherin was observed at the cell membrane, and relative expression level results revealed no significant relationship between the ECM sheet and the control. In contrast, β-catenin was densely localized at the cell perimeters and the cell interiors around the cell nuclei on the ECM sheet. The expression of βcatenin revealed significantly higher levels on the ECM sheet than on the control as shown by the staining results. The HUVEC adherens junctions on the ECM sheet, composed of various and complex ECM proteins, stimulated the matrix receptors of endothelial cells.30,31 VE-cadherin constitutes an extracellular domain, which is a junction responsible for calcium-dependent cell−cell adhesion of endothelial cells.32 It has long been recognized that interstitial ECMs, such as collagen I and fibronectin/fibrin, not only contribute to the formation of actin stress fibers but also disrupt VE-cadherin from intercellular junctions, thus facilitating multicellular reorganization.33 The relatively weak and disrupted VE-cadherin may be associated with collagen- and fibronectin-rich components of the ECM sheet. Meanwhile, βcatenin plays an important role in the regulation of vascular endothelial cell−cell adhesions.29 In particular, the β-catenin is an intracellular junction protein that interacts with the assembly of F-actin.34 Thus, the localized β-catenin in the cell interiors on the ECM sheet may have resulted from that the binding of β-catenin to distributed actin filaments. Collectively, our results indicate that physical and biochemical cues derived from the ECM sheet could differentially regulate endothelial cell−cell interaction in an orchestrated manner. Shear Stress-Induced Focal Adhesion and Adherens Junction Reorganization. We further examined whether the endothelial cellular response to altered shear stress can be regulated by culture conditions. Shear stress was applied to the confluent recellularized endothelial cells at 0.5, 1, and 5 dyn/ cm2 for 2 h. The tested shear stress is similar to the human blood vessel conditions; the venous shear stress ranges from 1 to 6 dyn/cm2.35 Shear stress-induced mechanical stimulation enhanced the polymerization of focal adhesion on the ECM sheet, as shown in Figure 5. Under low-shear stress conditions (0.5 dyn/cm2), vinculins assembled into dot and short-line formations in the cell interiors and at the cell membrane. At 5 dyn/cm2, mature vinculin sites were observed in long, thin lines. In contrast, on the control surface, vinculin fibers were not polymerized at high shear stress condition. The VEcadherin expression pattern was markedly different from that of the control. On ECM sheet, the VE-cadherin was depolymerized as fluid shear stress increased; VE-cadherin was localized, short, and thick at 0.5 dyn/cm2, but localized, long, and thin at the 5 dyn/cm2. It is known that VE-cadherin could recognize the junction in response to an applied forces, recruiting additional cadherin molecules to the junctions and vinculin to strengthen the link between catenin and actin. Accordingly, our results indicate that long lines of vinculin were correlated with VE-cadherin reorganization. These results suggest that endothelial cell response to shear stress is dependent on the ECM microenvironment. These findings can be applied to advance the engineering of vascular-associated disease models.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Kwideok Park: 0000-0001-8519-8316 Pilnam Kim: 0000-0003-0611-6525 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This research supported by the National Research Foundation of Korea (NRF) (grant number: NRF2016R1E1A1A01943393) and from the Ministry of Science ICT and Future Planning of Korea (grant number: 2016M3A9B4915823). We also appreciate the financial support from the R&D convergence Program of NST (National Research council of Science & Technology of Republic of Korea (grant number: CAP-14-3-KRISS).

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REFERENCES

(1) Frey, O.; Misun, P. M.; Fluri, D. A.; Hengstler, J. G.; Hierlemann, A. Reconfigurable microfluidic hanging drop network for multi-tissue interaction and analysis. Nat. Commun. 2014, 5, 4250 and Supporting Information therein. (2) Jang, M.; Koh, I.; Lee, S. J.; Cheong, J.-H.; Kim, P. Droplet-based microtumor model to assess cell-ECM interactions and drug resistance of gastric cancer cells. Sci. Rep. 2017, 7, 41541 and Supporting Information therein. (3) Jang, M.; Yang, S.; Kim, P. Microdroplet-based cell culture models and their application. BioChip J. 2016, 10 (4), 310−317. (4) Wang, T.; Zhang, M.; Dreher, D. D.; Zeng, Y. Ultrasensitive microfluidic solid-phase ELISA using an actuatable microwellpatterned PDMS chip. Lab Chip 2013, 13 (21), 4190−4197. (5) Gerver, R. E.; Herr, A. E. Microfluidic Western Blotting of LowMolecular-Mass Proteins. Anal. Chem. 2014, 86 (21), 10625−10632. (6) Blundell, C.; Tess, E. R.; Schanzer, A. S. R.; Coutifaris, C.; Su, E. J.; Parry, S.; Huh, D. A microphysiological model of the human placental barrier. Lab Chip 2016, 16 (16), 3065−3073. (7) van der Helm, M. W.; van der Meer, A. D.; Eijkel, J. C. T.; van den Berg, A.; Segerink, L. I. Microfluidic organ-on-chip technology for blood-brain barrier research. Tissue Barriers 2016, 4 (1), e1142493. (8) Zhang, Y. S.; Aleman, J.; Arneri, A.; Bersini, S.; Piraino, F.; Shin, S. R.; Dokmeci, M. R.; Khademhosseini, A. From cardiac tissue engineering to heart-on-a-chip: beating challenges. Biomed. Mater. 2015, 10 (3), 034006.

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DOI: 10.1021/acsbiomaterials.7b00613 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering (9) Aung, A.; Theprungsirikul, J.; Lim, H. L.; Varghese, S. Chemotaxis-driven assembly of endothelial barrier in a tumor-on-achip platform. Lab Chip 2016, 16 (10), 1886−98. (10) van Duinen, V.; Trietsch, S. J.; Joore, J.; Vulto, P.; Hankemeier, T. Microfluidic 3D cell culture: from tools to tissue models. Curr. Opin. Biotechnol. 2015, 35, 118−126. (11) Michielin, F.; Serena, E.; Pavan, P.; Elvassore, N. Microfluidicassisted cyclic mechanical stimulation affects cellular membrane integrity in a human muscular dystrophy in vitro model. RSC Adv. 2015, 5 (119), 98429−98439. (12) Jang, K.-J.; Suh, K.-Y. A multi-layer microfluidic device for efficient culture and analysis of renal tubular cells. Lab Chip 2010, 10 (1), 36−42. (13) Agarwal, A.; Goss, J. A.; Cho, A.; McCain, M. L.; Parker, K. K. Microfluidic heart on a chip for higher throughput pharmacological studies. Lab Chip 2013, 13 (18), 3599−3608. (14) Wang, X.-Y.; Fillafer, C.; Pichl, C.; Deinhammer, S.; HoferWarbinek, R.; Wirth, M.; Gabor, F. A multichannel acoustically driven microfluidic chip to study particle-cell interactions. Biomicrofluidics 2013, 7 (4), 044127. (15) Yang, K.; Lee, J. S.; Kim, J.; Lee, Y. B.; Shin, H.; Um, S. H.; Kim, J. B.; Park, K. I.; Lee, H.; Cho, S.-W. Polydopamine-mediated surface modification of scaffold materials for human neural stem cell engineering. Biomaterials 2012, 33 (29), 6952−6964. (16) Wang, L.; Sun, B.; Ziemer, K. S.; Barabino, G. A.; Carrier, R. L. Chemical and physical modifications to poly(dimethylsiloxane) surfaces affect adhesion of Caco-2 cells. J. Biomed. Mater. Res., Part A 2010, 93 (4), 1260−1271. (17) Kim, H. D.; Lee, E. A.; An, Y.-H.; Kim, S. L.; Lee, S. S.; Yu, S. J.; Jang, H. L.; Nam, K. T.; Im, S. G.; Hwang, N. S. Chondroitin SulfateBased Biomineralizing Surface Hydrogels for Bone Tissue Engineering. ACS Appl. Mater. Interfaces 2017, 9 (26), 21639−21650. (18) Decaris, M. L.; Binder, B. Y.; Soicher, M. A.; Bhat, A.; Leach, J. K. Cell-derived matrix coatings for polymeric scaffolds. Tissue Eng., Part A 2012, 18 (19−20), 2148−57. (19) Cheng, H. W.; Tsui, Y. K.; Cheung, K. M.; Chan, D.; Chan, B. P. Decellularization of chondrocyte-encapsulated collagen microspheres: a three-dimensional model to study the effects of acellular matrix on stem cell fate. Tissue Eng., Part C 2009, 15 (4), 697−706. (20) Song, J. J.; Guyette, J.; Gilpin, S.; Gonzalez, G.; Vacanti, J. P.; Ott, H. C. Regeneration and Experimental Orthotopic Transplantation of a Bioengineered Kidney. Nat. Med. 2013, 19 (5), 646−651. (21) Xing, Q.; Qian, Z.; Jia, W.; Ghosh, A.; Tahtinen, M.; Zhao, F. Natural Extracellular Matrix for Cellular and Tissue Biomanufacturing. ACS Biomater. Sci. Eng. 2017, 3 (8), 1462−1476. (22) Park, G. R.; Lee, J. G.; Chun, H. J.; Han, D. K.; Park, K. Characterization of naturally derived macromolecular matrix and its osteogenic activity with preosteoblasts. Macromol. Res. 2012, 20 (8), 868−874. (23) Xing, Q.; Yates, K.; Tahtinen, M.; Shearier, E.; Qian, Z.; Zhao, F. Decellularization of fibroblast cell sheets for natural extracellular matrix scaffold preparation. Tissue Eng., Part C 2015, 21 (1), 77−87. (24) Lee, J. S.; Lee, K.; Moon, S.-H.; Chung, H.-M.; Lee, J. H.; Um, S. H.; Kim, D.-I.; Cho, S.-W. Mussel-Inspired Cell-Adhesion Peptide Modification for Enhanced Endothelialization of Decellularized Blood Vessels. Macromol. Biosci. 2014, 14 (8), 1181−1189. (25) Zhang, W.; Zhu, Y.; Li, J.; Guo, Q.; Peng, J.; Liu, S.; Yang, J.; Wang, Y. Cell-Derived Extracellular Matrix: Basic Characteristics and Current Applications in Orthopedic Tissue Engineering. Tissue Eng., Part B 2016, 22 (3), 193−207. (26) Fadini, G. P.; Avogaro, A. Cell-based methods for ex vivo evaluation of human endothelial biology. Cardiovasc. Res. 2010, 87 (1), 12−21. (27) Cukierman, E.; Pankov, R.; Stevens, D. R.; Yamada, K. M. Taking Cell-Matrix Adhesions to the Third Dimension. Science 2001, 294 (5547), 1708−1712. (28) Petrie, R. J.; Doyle, A. D.; Yamada, K. M. Random versus directionally persistent cell migration. Nat. Rev. Mol. Cell Biol. 2009, 10 (8), 538−549.

(29) Lampugnani, M. G.; Corada, M.; Caveda, L.; Breviario, F.; Ayalon, O.; Geiger, B.; Dejana, E. The molecular organization of endothelial cell to cell junctions: differential association of plakoglobin, beta-catenin, and alpha-catenin with vascular endothelial cadherin (VE-cadherin). J. Cell Biol. 1995, 129 (1), 203−217. (30) Baeten, K. M.; Akassoglou, K. Extracellular Matrix and Matrix Receptors in Blood-Brain Barrier Formation and Stroke. Dev. Neurobiol. 2011, 71 (11), 1018−1039. (31) Cleaver, O.; Melton, D. A. Endothelial signaling during development. Nat. Med. 2003, 9 (6), 661−668 and Supporting Information therein. (32) Herren, B.; Levkau, B.; Raines, E. W.; Ross, R. Cleavage of betacatenin and plakoglobin and shedding of VE-cadherin during endothelial apoptosis: evidence for a role for caspases and metalloproteinases. Mol. Biol. Cell 1998, 9 (6), 1589−601. (33) Davis, G. E.; Senger, D. R. Endothelial Extracellular Matrix: Biosynthesis, Remodeling, and Functions During Vascular Morphogenesis and Neovessel Stabilization. Circ. Res. 2005, 97 (11), 1093− 1107. (34) Drees, F.; Pokutta, S.; Yamada, S.; Nelson, W. J.; Weis, W. I. αCatenin Is a Molecular Switch that Binds E-Cadherin-β-Catenin and Regulates Actin-Filament Assembly. Cell 2005, 123 (5), 903−915. (35) Malek, A. M.; Alper, S. L.; Izumo, S. Hemodynamic shear stress and its role in atherosclerosis. JAMA 1999, 282 (21), 2035−2042.

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DOI: 10.1021/acsbiomaterials.7b00613 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX